We acknowledge funding ARO MURI W911NF-14-1-0403 to M.P.M., the National Institutes of Health (NIH) R01 1R01GM126256 to M.P.M., the National Institutes of Health (NIH) U54 CA209992, NIH RO1 GM126256, NIH U54 CA209992, University of Michigan / Genentech, SUBK00016255 and Human Frontiers Science Program (HFSP) grant number RGY0073/2018 to M.P.M. Any opinion, findings, conclusions, or recommendations expressed in this material are those of the authors(s) and do not necessarily reflect the views of the ARO, NIH, or HFSP. S.C. acknowledges fruitful discussions with V. Yadav, C. Muresan, and S. Amiri.

1. Preparation of buffers and protein solutions
<ol>
	<li>Prepare the aqueous Inner Non-Polymerization (INP) Buffer in a total volume of 5 mL by mixing 0.1 mM CaCl2, 10 mM HEPES (pH 7.5), 1 mM DTT, 0.5 mM Dabco, 320 mM sucrose, and 0.2 mM ATP.</li>
	<li>Prepare the Protein Mix (PM) by adding proteins to the INP buffer at 4 &#176;C with the following concentrations: 11.2 &#181;M non-fluorescent G-actin, 2.8 &#181;M fluorescently labeled actin, and 0.24 &#181;M Arp2/3 (Table of Materials). To form an F-actin layer, add 100 nM gelsolin, 4 &#181;M cofilin, and 2.2 &#181;M VCA-His to the PM. As a control experiment, replace PM by 100 &#181;g/mL fluorescent dye (Table of Materials).</li>
	<li>Prepare the aqueous Inner Polymerization (IP) Buffer (aqueous) in a total volume of 5 mL by mixing 100 mM KCl, 4 mM MgCl2, 10 mM HEPES (pH 7.5), 1 mM DTT, 0.5 mM Dabco, 10 mM ATP, and 80 mM sucrose.</li>
	<li>Prepare the Final Buffer (FB) by mixing INP buffer (containing PM) and IP buffer at a volume ratio of 1:1 to yield the inner aqueous solution in a total volume of 30 &#181;L that will be encapsulated within the liposome.</li>
	<li>Prepare the aqueous Outside Buffer (OB) in a total volume of 150 &#181;L by mixing 10 mM HEPES (pH 7.5), 50 mM KCl, 2mM MgCl2, 0.2 mM CaCl2, 2mM ATP, 1 mM DTT, 0.5 mM Dabco, 212 mM glucose, and 0.1 mg/mL &#946;-casein. 
	NOTE: The osmolarity of the OB can be adjusted with glucose to ensure the osmotic pressure of the OB is slightly larger than that of the FB (20-60 mOsm). The density of the FB should be slightly higher than the OB.</li>
</ol>
2. Preparation of liposomes based on the inverted emulsion techniques
<ol>
	<li>Prepare the lipid-oil mixture
	<ol>
		<li>Add 100 &#181;L of 25 mg/mL L-&#945;-phosphatidylcholine (non-fluorescent Egg PC, also called EPC, including 1% DHPE) into a glass vial. Evaporate chloroform with argon gas, leaving a dry solid lipid film (2.5 mg) at the bottom of the vial.
		<ol>
			<li>To form an F-actin layer, add 1,2-dioleoyl-sn-glycero-3-{[n(5-amino-1-carboxypentyl) iminodiacetic acid] succinyl} nickel salt (DOGS-NTA-Ni) at a 10:1 ratio of EPC to DOGS-NTA-Ni and mix before the evaporation of chloroform.</li>
		</ol>
		</li>
		<li>Add 2 mL of mineral oil and sonicate the lipid-oil mixture at room temperature in a bath for 1 h to re-suspend the lipids. 
		NOTE: The lipid-oil mixture after sonication can be kept for 1 week at 4 &#176;C. Re-sonication is suggested before use.</li>
	</ol>
	</li>
	<li>Prepare a Final Buffer in oil (FB/oil) emulsion for preparing monolayer lipid droplets containing the protein of interest.
	<ol>
		<li>To 100 &#181;L of lipid-oil mixture taken in a plastic tube, add 10 &#181;L of FB. Ensure that FB is in one droplet.</li>
		<li>Using a glass syringe, draw the lipid-oil-FB mixture and gently aspirate up and down multiple times to emulsify it; draw a small amount of lipid-oil mixture first, and then the FB droplet by placing the tip of the syringe at the periphery of the droplet to break it up into tiny droplets. Repeat the up and down aspiration until a whitish and cloudy emulsion is formed.</li>
	</ol>
	</li>
	<li>Put 30 &#181;L of OB in a separate plastic tube. Place 30 &#181;L of the lipid-oil mixture on top of OB and let it sit for ~10 mins to develop a lipid monolayer at the interface. 
	NOTE: If the lipids are charged or the protein is incorporated, the duration of this step should be extended38.</li>
	<li>Prepare the liposomes
	<ol>
		<li>Carefully add 50 &#181;L of the FB/oil emulsion (step 2.2) to the top oil phase of the tube from step 2.3.</li>
		<li>Centrifuge the plastic tube at 100 x g for 15 mins at 4 &#176;C. Vary time and centrifugation speed to optimize for liposome formation. 
		NOTE: After centrifugation, the top oil phase should be clear, and the bottom OB (containing liposomes) should be slightly cloudy.</li>
		<li>Carefully remove the oil phase away by pipette. Aspirate extra volume if needed. Ensure not to put the pipette tip at the side of the tube to avoid creating a meniscus of oil on top of the liposome phase.</li>
		<li>With a new pipette, slowly stick the pipette tip into the remaining bottom phase and aspirate the aqueous volume to collect liposomes. 
		NOTE: It is better to lose volume than to incorporate the oil. The inclusion of oil will cause liposomes to rupture, and the internal components will be released into the external buffer. Cut the tip of a pipette to reduce shear.</li>
	</ol>
	</li>
</ol>
3. Microscopy observation
<ol>
	<li>Pour 100 &#181;L of OB into the well of an incubation chamber (4-well plate containing 12 mm round coverslips). Gently deposit the collected liposomes (step 2.4.4) into OB, and then place another coverslip on top of the chamber. 
	NOTE: The OB contains the &#946;-casein to passivate the glass surface and minimize sticking between the liposomes20.</li>
	<li>Observe liposomes with a confocal microscope using a 63x oil-immersion objective. Use 488 nm, 647 nm, and 561 nm laser lines to observe fluorescently labeled lipid, encapsulated fluorescent dye, and encapsulated fluorescently labeled actin, respectively. Capture the frames of interest and save the images in TIFF format.</li>
	<li>Process and analyze images in ImageJ/Fiji39. For new users of ImageJ/Fiji, an online tutorial is available (see Table of Materials).
	<ol>
		<li>Open the TIFF file using ImageJ/Fiji software.</li>
		<li>Go to Image &#62; Adjust &#62; Brightness/Contrast. Adjust the brightness and contrast of the image to the desired scale by adjusting the Minimum and Maximum settings, and the Brightness, and Contrast sliders.</li>
		<li>Click on the Rectangle selection tool in the toolbar and select the region of interest (ROI). Go to Image &#62; Crop to crop the ROI.</li>
		<li>Go to Analyze &#62; Set Scale. In the pop-up window, enter 1.0&#160;in the Pixel Aspect Ratio field and set &#956;m as the Unit of Length. In the Distance in Pixels field, enter the image width in pixels. In the Known Distance field, enter the real image width in microns.</li>
		<li>To measure the size of the liposome, Using the Oval selection tool in the toolbar, draw a circle along the edge of the liposome. Go to Analyze &#62; Measure to measure the area of the circle, from which the diameter of the liposome can be calculated.</li>
	</ol>
	</li>
</ol>

The authors declare no conflicts of interest.

Several key steps determine the success of a high yield of liposomes during the preparation process. To completely dissolve the lipid film in the oil, the sample must be sonicated until the lipid film at the bottom of the glass vial disappears completely. After the sonication, the lipid-oil mixture must be stored overnight at room temperature under dark conditions for the lipid molecules to disperse further29. The mixture can be stored at 4 &#176;C for up to a week. When preparing an FB/oil emulsi...

The actin cytoskeleton plays a fundamental role in constructing the intracellular architecture of the cell by coordinating molecular-level contractility and force generation1,2,3. As a result, it mediates numerous essential cellular activities, including cell deformation4,5, division6, migration7,8, and adhesion9. The in vitro reconstitution of actin networks has gained tremendous attention in recent years10,11,12,13,14,15,16,17. The goal of reconstitution is to build a minimal model of the cell devoid of the complex biochemical regulation that exists within live cells. This offers a controllable environment to probe specific intracellular activities and facilitates the identification and analysis of different components of the actin cytoskeleton18,19. Further, the encapsulation of in vitro actin networks inside phospholipid giant unilamellar vesicles (GUVs, liposomes) provides a confined but deformable space with a semi-permeable boundary. It mimics the physiological and mechanical microenvironment of the actin machinery within the cell9,20,21,22.
Among various methods to prepare liposomes, the lipid film hydration method (also known as the swelling method) is one of the earliest techniques23. The dry lipid film hydrates with the addition of buffers, forming membranous bubbles that eventually become vesicles24. To produce larger vesicles with a higher yield, an improved method advancing from the film hydration method, known as the electroformation method25, applies an AC electric field to efficiently promote the hydration process26. The major limitations of these hydration-based methods for actin encapsulation are that it has low encapsulation efficiency of highly concentrated proteins, and it is only compatible with specific lipid compositions24. The inverted emulsion technique, in comparison, has fewer limitations for lipid components and protein concentrations20,27,28,29. In this method, a mixture of proteins for encapsulation is added to the inner aqueous buffer, which is later emulsified in a lipid-containing mineral oil solution, forming lipid-monolayer droplets. The monolayer lipid droplets then cross through another lipid/oil-water interface through centrifugation to form bilayer lipid vesicles (liposomes). This technique has proven to be one of the most successful strategies for actin encapsulation24,30. Separately, there are some microfluidic device methods, including pulsed jetting31,32, transient membrane ejection33, and the cDICE method34. The similarities between the inverted emulsion method and the microfluidic method are the lipid solvent (oil) that is utilized and the introduction of lipid/oil-water interface for the formation of the outer leaflet of liposomes. By contrast, the generation of liposomes by the microfluidic method requires a set-up of microfluidic devices and is accompanied by oil trapped between the two leaflets of the bilayer, which requires an extra step for oil removal35.
In this manuscript, we used the inverted emulsion technique to prepare liposomes encapsulating a polymerized F-actin network as used previously22. The protein mixture for encapsulation was first placed in a buffer with nonpolymerizing conditions to maintain actin in its globular (G) form. The whole process was carried out at 4 &#176;C to prevent early actin polymerization, which was later triggered by allowing the sample to warm to room temperature. Once at room temperature, the actin polymerizes into its filamentous (F) form. A variety of actin-binding proteins can be added to the inner aqueous buffer solution to study protein functionalities and properties, thus, further providing insights into its interaction with the actin network and membrane surface. This method can also be applied to the encapsulation of various proteins of interest36&#160;and large objects (microparticles, self-propelled microswimmers, etc.) close to the size of the final liposomes28,37.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3-{[n(5-amino-1-carboxypentyl)iminodiacetic acid]succinyl} nickel salt (DOGS-NTA-Ni)&#160;</td>
			<td>Avanti Polar Lipids Inc.</td>
			<td>231615773</td>
			<td>Nickel Lipid</td>
		</tr>
		<tr>
			<td>1,4-Diazabicyclo[2.2.2]octane</td>
			<td>Sigma</td>
			<td>D27802-25G</td>
			<td>DABCO</td>
		</tr>
		<tr>
			<td>Actin protein (&#62;99% pure): rabbit skeletal muscle</td>
			<td>Cytoskeleton, Inc</td>
			<td>AKL99-D</td>
			<td>non-fluorescent G-actin</td>
		</tr>
		<tr>
			<td>Actin protein (rhodamine): rabbit skeletal muscle</td>
			<td>Cytoskeleton, Inc</td>
			<td>AR05</td>
			<td>fluorescently labeled actin</td>
		</tr>
		<tr>
			<td>Adenosine 5&#8242;-triphosphate disodium salt hydrate</td>
			<td>Sigma</td>
			<td>A2383-10G</td>
			<td>ATP</td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 dye</td>
			<td>ThermoFisher</td>
			<td></td>
			<td>fluorescent dye</td>
		</tr>
		<tr>
			<td>Andor iQ3</td>
			<td>Andor Technologies</td>
			<td></td>
			<td>control and acquisition software for confocal microscope</td>
		</tr>
		<tr>
			<td>Arp2/3 Protein Complex: Porcine Brain</td>
			<td>Cytoskeleton, Inc</td>
			<td>RP01P-A</td>
			<td>Arp 2/3</td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma</td>
			<td>10035048</td>
			<td>CaCl2</td>
		</tr>
		<tr>
			<td>Chamlide Chambers (4-well for 12 mm round coverslip)</td>
			<td>Quorum Technologies</td>
			<td></td>
			<td>incubation chamber</td>
		</tr>
		<tr>
			<td>Cofilin protein: human recombinant</td>
			<td>Cytoskeleton, Inc</td>
			<td>CF01-C</td>
			<td>cofilin</td>
		</tr>
		<tr>
			<td>Confocal Microscope (63&#215; oil-immersion objective)</td>
			<td>Andor Technologies</td>
			<td>LEICA DMi8</td>
			<td></td>
		</tr>
		<tr>
			<td>D-(+)-GLUCOSE BIOXTRA</td>
			<td>Sigma</td>
			<td>G7528</td>
			<td>glucose</td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>DOT Scientific&#160;</td>
			<td>DSD11000-10</td>
			<td>DTT</td>
		</tr>
		<tr>
			<td>Gelsolin Protein: Homo Sapiens Recombinant</td>
			<td>Cytoskeleton, Inc</td>
			<td>HPG6</td>
			<td>gelsolin</td>
		</tr>
		<tr>
			<td>Hamilton 1750 Gastight Syringe, 500 &#181;L, cemented needle, 22 G, 2&#34; conical tip</td>
			<td>Cole-Parmer</td>
			<td>UX-07940-53</td>
			<td>glass syringe</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>AmericanBio</td>
			<td>7365-45-9</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ/Fiji</td>
			<td></td>
			<td></td>
			<td>https://imagej.net/tutorials/</td>
		</tr>
		<tr>
			<td>L-alpha-Phosphatidylcholine</td>
			<td>Avanti Polar Lipids Inc.</td>
			<td>97281442</td>
			<td>EPC</td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>Sigma</td>
			<td>7786303</td>
			<td>MgCl2</td>
		</tr>
		<tr>
			<td>Mineral oil, BioReagent, for molecular biology, light oil</td>
			<td>Sigma</td>
			<td>8042475</td>
			<td>mineral oil</td>
		</tr>
		<tr>
			<td>N-WASP fragment WWA (aa400&#8211;501, VCA-His)</td>
			<td></td>
			<td></td>
			<td>VCA-His is purified using lab protocol. The protocol can be provided upon reasonable requests</td>
		</tr>
		<tr>
			<td>Oregon Green 488 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine (Oregon Green 488 DHPE)</td>
			<td>Thermo Fisher</td>
			<td>O12650</td>
			<td>DHPE</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma</td>
			<td>7447407</td>
			<td>KCl</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>57-50-1</td>
			<td>sucrose</td>
		</tr>
		<tr>
			<td>&#946;-Casein from bovine milk</td>
			<td>Sigma</td>
			<td>C6905-250MG</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vitro reconstitution of the actin cytoskeleton inside giant unilamellar vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

The preparation of liposomes based on the inverted emulsion technique is illustrated graphically and schematically in Figure 1.
First, empty (bare) liposomes (~5-50 &#181;m in diameter) that were composed of phospholipid (EPC) and fluorescent lipid (DHPE) were prepared. A bright, far-red fluorescent dye was encapsulated within bare liposomes as a control experiment. Whether a lipid monolayer has successfully formed in the peripheral of the droplet could be determi...

Watch this Scientific Journal Video about In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles at JoVE.com

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell ...

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3.2K Views.  Yale University. In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

Video: In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Decellularization of Whole Human Heart Inside a Pressurized Pouch in an Inverted Orientation

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ultimately rejection can occur. Creating a functional, autologous bio-artificial heart could solve these challenges. Biofabrication of a heart comprised of scaffold and cells is one option. A natural scaffold with tissue-specific composition as well as micro- and macro-architecture can be obtained by decellularizing hearts from humans or large animals such as pigs. Decellularization involves washing out cellular debris while preserving 3D extracellular matrix and vasculature and allowing &#34;cellularization&#34; at a later timepoint. Capitalizing on our novel finding that perfusion decellularization of complex organs is possible, we developed a more &#34;physiological&#34; method to decellularize non-transplantable human hearts by placing them inside a pressurized pouch, in an inverted orientation, under controlled pressure. The purpose of using a pressurized pouch is to create pressure gradients across the aortic valve to keep it closed and improve myocardial perfusion. Simultaneous assessment of flow dynamics and cellular debris removal during decellularization allowed us to monitor both fluid inflow and debris outflow, thereby generating a scaffold that can be used either for simple cardiac repair (e.g. as a patch or valve scaffold) or as a whole-organ scaffold.

This method enables decellularization of a complex solid organ using a simple protocol based on osmotic shock and perfusion of ionic detergent with minimal organ matrix disruption. It comprises a novel decellularization technique for human hearts inside a pressurized pouch with real-time monitoring of flow dynamics and cellular debris outflow.

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ...

Advanced 3D Liver Models for In vitro Genotoxicity Testing Following Long-Term Nanomaterial Exposure

Due to the rapid development and implementation of a diverse array of engineered nanomaterials (ENM), exposure to ENM is inevitable and the development of robust, predictive in vitro test systems is essential. Hepatic toxicology is key when considering ENM exposure, as the liver serves a vital role in metabolic homeostasis and detoxification as well as being a major site of ENM accumulation post exposure. Based upon this and the accepted understanding that 2D hepatocyte models do not accurately mimic the complexities of intricate multi-cellular interactions and metabolic activity observed in vivo, there is a greater focus on the development of physiologically relevant 3D liver models tailored for ENM hazard assessment purposes in vitro. In line with the principles of the 3Rs to replace, reduce and refine animal experimentation, a 3D HepG2 cell-line based liver model has been developed, which is a user friendly, cost effective system that can support both extended and repeated ENM exposure regimes (&#8804;14 days). These spheroid models (&#8805;500 &#181;m in diameter) retain their proliferative capacity (i.e., dividing cell models) allowing them to be coupled with the &#8216;gold standard&#8217; micronucleus assay to effectively assess genotoxicity in vitro. Their ability to report on a range of toxicological endpoints (e.g., liver function, (pro-)inflammatory response, cytotoxicity and genotoxicity) has been characterized using several ENMs across both acute (24 h) and long-term (120 h) exposure regimes. This 3D in vitro hepatic model has the capacity to be utilized for evaluating more realistic ENM exposures, thereby providing a future in vitro approach to better support ENM hazard assessment in a routine and easily accessible manner.

This procedure was established to be used for developing advanced 3D hepatic cultures in vitro, which can provide a more physiologically relevant assessment of the genotoxic hazards associated with nanomaterial exposures over both an acute or long-term, repeated dose regimes.

Due to the rapid development and implementation of a diverse array of engineered nanomaterials (ENM), exposure to ENM is inevitable and the development of ...

An In vitro System to Gauge the Thrombolytic Efficacy of Histotripsy and a Lytic Drug

Deep vein thrombosis (DVT) is a global health concern. The primary approach to achieve vessel recanalization for critical obstructions is catheter-directed thrombolytics (CDT). To mitigate caustic side effects and the long treatment time associated with CDT, adjuvant and alternative approaches are under development. One such approach is histotripsy, a focused ultrasound therapy to ablate tissue via bubble cloud nucleation. Pre-clinical studies have demonstrated strong synergy between histotripsy and thrombolytics for clot degradation. This report outlines a benchtop method to assess the efficacy of histotripsy-aided thrombolytic therapy, or lysotripsy.
Clots manufactured from fresh human venous blood were introduced into a flow channel whose dimensions and acousto-mechanical properties mimic an iliofemoral vein. The channel was perfused with plasma and the lytic&#160;recombinant tissue-type plasminogen activator. Bubble clouds were generated in the clot with a focused ultrasound source designed for the treatment of femoral venous clots. Motorized positioners were used to translate the source focus along the clot length. At each insonation location, acoustic emissions from the bubble cloud were passively recorded, and beamformed to generate passive cavitation images. Metrics to gauge treatment efficacy included clot mass loss (overall treatment efficacy), and the concentrations of D-dimer (fibrinolysis) and hemoglobin (hemolysis) in the perfusate. There are limitations to this in vitro design, including lack of means to assess in vivo side effects or dynamic changes in flow rate as the clot lyses. Overall, the setup provides an effective method to assess the efficacy of histotripsy-based strategies to treat DVT.

Histotripsy-aided lytic delivery or lysotripsy is under development for the treatment of deep vein thrombosis. An in vitro procedure is presented here to assess the efficacy of this combination therapy. Key protocols for the clot model, image guidance, and assessment of treatment efficacy are discussed.

Deep vein thrombosis (DVT) is a global health concern. The primary approach to achieve vessel recanalization for critical obstructions is ...

Labeling of Extracellular Vesicles for Monitoring Migration and Uptake in Cartilage Explants

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their cellular source, such as platelet lysate (PL). When used as treatment, EVs are expected to enter the target cells and effect a response from these. In this research, PL-derived EVs have been studied as a cell-free treatment for osteoarthritis (OA). Thus, a method was set up to label EVs and test their uptake on cartilage explants. PL-derived EVs are labeled with the lipophilic dye PKH26, washed twice through a column, and then tested in an in vitro inflammation-driven&#160;OA model for 5 h after particle quantification by nanoparticle tracking analysis (NTA). Hourly, cartilage explants are fixed, paraffined, cut into 6 &#181;m sections to mount on slides, and observed under a confocal microscope. This allows verification of whether EVs enter the target cells (chondrocytes) during this period and analyze their direct effect.

Here, we present a protocol to label platelet lysate-derived extracellular vesicles to monitor their migration and uptake in cartilage explants used as a model for osteoarthritis.

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their ...

Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular processes in vitro. In recent years, encapsulation within GUVs has proven to be a helpful approach for reconstitution experiments in cell biology and related fields. It better mimics confinement conditions inside living cells, as opposed to conventional biochemical reconstitution. Methods for encapsulation inside GUVs are often not easy to implement, and success rates can differ significantly from lab to lab. One technique that has proven to be successful for encapsulating more complex protein systems is called continuous droplet interface crossing encapsulation (cDICE). Here, a cDICE-based method is presented for rapidly encapsulating cytoskeletal proteins in GUVs with high encapsulation efficiency. In this method, first, lipid-monolayer droplets are generated by emulsifying a protein solution of interest in a lipid/oil mixture. After being added into a rotating 3D-printed chamber, these lipid-monolayered droplets then pass through a second lipid monolayer at a water/oil interface inside the chamber to form GUVs that contain the protein system. This method simplifies the overall procedure of encapsulation within GUVs and speeds up the process, and thus allows us to confine and observe the dynamic evolution of network assembly inside lipid bilayer vesicles. This platform is handy for studying the mechanics of cytoskeleton-membrane interactions in confinement.

This article introduces a simple method for expeditious production of giant unilamellar vesicles with encapsulated cytoskeletal proteins. The method proves to be useful for bottom-up reconstitution of cytoskeletal structures in confinement and cytoskeleton-membrane interactions.

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular ...

Reconstitution of Actin-Based Motility with Commercially Available Proteins

Many cell movements and shape changes and certain types of intracellular bacterial and organelle motility are driven by the biopolymer actin that forms a dynamic network at the surface of the cell, organelle, or bacterium. The biochemical and mechanical basis of force production during this process can be studied by reproducing actin-based movement in an acellular manner on inert surfaces such as beads that are functionalized and incubated with a controlled set of components. Under the appropriate conditions, an elastic actin network assembles at the bead surface and breaks open due to the stress generated by network growth, forming an "actin comet" that propels the bead forward. However, such experiments require the purification of a host of different actin-binding proteins, often putting them beyond the reach of non-specialists. This article details a protocol for reproducibly obtaining actin comets and motility of beads using commercially available reagents. Bead coating, bead size, and motility mixture can be altered to observe the effect on bead speed, trajectories, and other parameters. This assay can be used for testing the biochemical activities of different actin-binding proteins, and for performing quantitative physical measurements that shed light on active matter properties of actin networks. This will be a useful tool for the community, enabling the study of in vitro actin-based motility without expert knowledge in actin-binding protein purification.

This protocol describes how to produce actin comets on the surfaces of beads using commercially available protein ingredients. Such systems mimic the protrusive structures found in cells, and can be used to examine physiological mechanisms of force production in a simplified way.

Many cell movements and shape changes and certain types of intracellular bacterial and organelle motility are driven by the biopolymer actin that forms a ...

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This feature is attributed to the mechanical and dynamic properties of the cell cytoskeleton, notably, the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin motors, and accessory proteins that exhibit intrinsic contraction properties. The usually accepted view is that the cytoskeleton behaves as a viscoelastic material. However, this model cannot always explain the experimental results, which are more consistent with a picture describing the cytoskeleton as a poroelastic active material-an elastic network embedded with cytosol. Contractility gradients generated by the myosin motors drive the flow of the cytosol across the gel pores, which infers that the mechanics of the cytoskeleton and the cytosol are tightly coupled. One main feature of poroelasticity is the diffusive relaxation of stresses in the network, characterized by an effective diffusion constant that depends on the gel elastic modulus, porosity, and cytosol (solvent) viscosity. As cells have many ways to regulate their structure and material properties, our current understanding of how cytoskeleton mechanics and cytosol flow dynamics are coupled remains poorly understood. Here, an in vitro reconstitution approach is employed to characterize the material properties of poroelastic actomyosin gels as a model system for the cell cytoskeleton. Gel contraction is driven by myosin motor contractility, which leads to the emergence of a flow of the penetrating solvent. The paper describes how to prepare these gels and run experiments. We also discuss how to measure and analyze the solvent flow and gel contraction both at the local and global scales. The various scaling relations used for data quantification are given. Finally, the experimental challenges and common pitfalls are discussed, including their relevance to cell cytoskeleton mechanics.

The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

In this work, an in vitro reconstitution approach is employed to study the poroelasticity of actomyosin gels under controlled conditions. The dynamics of the actomyosin gel and the embedded solvent are quantified, through which the network poroelasticity is demonstrated. We also discuss the experimental challenges, common pitfalls, and relevance to cell cytoskeleton mechanics.

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This ...

Light-Controlled Reversible Protein Patterning on Lipid Membranes

Dynamic Light-Induced Protein Patterns at Model Membranes

The precise localization and activation of proteins at the cell membrane at a certain time gives rise to many cellular processes, including cell polarization, migration, and division. Thus, methods to recruit proteins to model membranes with subcellular resolution and high temporal control are essential when reproducing and controlling such processes in synthetic cells. Here, a method is described for fabricating light-regulated reversible protein patterns at lipid membranes with high spatiotemporal precision. For this purpose, we immobilize the photoswitchable protein iLID (improved light-inducible dimer) on supported lipid bilayers (SLBs) and on the outer membrane of giant unilamellar vesicles (GUVs). Upon local blue light illumination, iLID binds to its partner Nano (wild-type SspB) and allows the recruitment of any protein of interest (POI) fused to Nano from the solution to the illuminated area on the membrane. This binding is reversible in the dark, which provides dynamic binding and release of the POI. Overall, this is a flexible and versatile method for regulating the localization of proteins with high precision in space and time using blue light.

Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems

Here, a protocol is described for generating light-regulated and reversible protein patterns with high spatiotemporal precision at artificial lipid membranes. The method consists of the localized photoactivation of the protein iLID (improved light-inducible dimer) immobilized on model membranes that, under blue light, binds to its partner protein Nano (wild-type SspB).